Submitted to the
Combined Faculties for the Natural Sciences and for Mathematics
Of the Ruperto-Carola University of Heidelberg, Germany
for the degree of
Doctor of Natural Sciences
presented by
Diplom-Biology: Yaodong Yang
Born in: Xiamen, China
From the Endoplasmic Reticulum to the Golgi Apparatus:
In Vitro and In Vivo Approaches to
Understanding COPII Vesicle Function in Plant Cells
Referees:
Prof.
Dr.
David
G.
Robinson
TABLE OF CONTENTS...I SUMMARY...IV
1. INTRODUCTION... 1
1.1. An overview of the secretory pathway ... 1
1.2. Secretion-related Organelles and Compartments in Plant Cells ... 2
1.2.1. Endoplasmic Reticulum (ER) ... 2
1.2.2. Golgi Apparatus (GA)... 5
1.2.3. Prevacuolar Compartment (PVC) ... 7
1.2.4. Vacuoles... 8
1.3. Vesicles mediating cargo transport in the early secretory pathway... 9
1.3.1. COPI Vesicles... 10
1.3.2. COPII Vesicles... 13
1.3.2.1. COPII coat components ... 14
1.3.2.2. COPII formation and disassembly ... 16
1.3.2.3. Protein Sorting into COPII vesicles... 19
1.3.2.4. COPII vesicles in plants... 21
1.4. SNAREs and Membrane Fusion... 22
1.4.1. SNAREs... 22 1.4.2. Other proteins... 24 1.4.2.1. SM Proteins... 24 1.4.2.2. Rab Proteins ... 25 1.4.2.3. Tethering Factors ... 26 1.5. Objectives ... 27
2. MATERIALS and METHODS ... 28
2.1. Culture media... 28
2.1.1. Arabidopsis culture media: ... 28
2.1.1.1. Seed germination media (1/2MS) ... 28
2.1.1.2. Callus induction medium and culture media ... 28
2.1.2. Tobacco Bright Yellow 2 (BY-2) culture medium:... 28
2.1.3. Bacteria culture meium:... 28
2.2. Organisms and Culture Techniques ... 29
2.2.1. Bacteria ... 29
2.2.2. Arabidopsis ... 29
2.2.2.1. Seed germination ... 29
2.2.2.2. Callus induction ... 29
2.2.2.3. Preparation of cell suspension cultures... 30
2.2.3. BY-2 cell suspension culture ... 30
2.3 Molecular Biological Methods ... 30
2.3.1. Bacterial plasmid DNA preparation... 30
2.3.1.1. Minipreps of plasmid DNA ... 30
2.3.1.2. Midipreps of plasmid DNA ... 30
2.3.2. Preparing samples for sequencing ... 31
2.3.3. Spectrophotometric estimation of DNA purity and quantitation... 31
2.3.4. Enzymatic treatment of DNA ... 31
2.3.5. Transformation of E. coli... 32
2.3.5.2. Preparation of competent cells and transformation by electroporation ... 33
2.3.6 Transformation of Agrobacterium tumefaciens ... 33
2.3.7 BY-2 Stable Transfromation... 34
2.3.8 PCR amplification of DNA... 34
2.3.9 Plasmid constructions ... 35
2.4 Biochemical Methods ... 38
2.4.1 Polyacrylamide gel electrophoresis (PAGE) ... 38
2.4.2. Preparative gel electrophoresis and electro-elution ... 40
2.4.3. Western blotting and immunological detection of proteins on nitrocellulose filters ... 40
2.4.4. Protein quantitation... 41
2.4.5. Concentrating proteins ... 41
2.4.6. Protein expression and purification ... 41
2.4.6.1. Protein extraction from bacteria... 42
2.4.6.2. Protein purification ... 42
2.4.7. Antibody purification... 46
2.4.8. Subcellular fractionation... 47
2.4.8.1. Preparation of cytosol and microsomal fractions... 47
2.4.8.2. Sucrose-density-gradient centrifugation ... 47
2.4.8.3. Flotation gradient:... 47
2.4.9. COPII In vitro budding assay: ... 48
2.4.9.1. Preparation of 30% (NH4)2SO4 cut off cytosol:... 48
2.4.9.2. Preparation of ER-enriched microsomes: ... 48
2.4.9.3. Budding conditions ... 48
2.4.10. In vitro binding of COPI and COPII coats to sorting motifs in p24 proteins 49 2.5. Microscopical analysis... 50
2.5.1 Confocal Laser Scanning Microscopy ... 50
2.5.1.1. Immunofluorescence Labeling... 50
2.5.1.2. Microscopy ... 51
2.5.2. Electron microscopy ... 51
2.5.2.1 Negative staining: ... 51
2.5.2.2. Immunogold Negative Staining ... 51
2.6. DNA and protein sequence computer analysis ... 52
3. RESULTS ... 53
3.1. Searching for COPII related proteins in Arabidopsis ... 53
3.1.1 Cloning and expression fusion proteins... 54
3.1.2. Antibodies ... 55
3.1.3. The principal subcellular location of the proteins... 56
3.2. In vitro approach:... 57
3.2.1. Yeast COPII vesicle budding... 57
3.2.2. COPII proteins salt out with 30% ammonium sulphate... 58
3.2.3. The integrity of AtSec13 and AtSec23 complexes ... 59
3.2.4. COPII vesicle budding with Arabidopsis ... 60
3.2.5. Factors affecting budding ... 61
3.2.6. Purification of putative COPII vesicle... 64
3.3. In vivo approach: ... 73
3.3.1. AtSec13 and AtSar1p are presnet on the ER of Arabidopsis... 73
3.3.2. COPII immunostaining in BY-2 cell lines expressing fluorescent ER and Golgi markers... 74
3.3.3 Establishment of a BY-2 cell line expressing Sec13-GFP... 77
3.3.4. ERESs in relation to the ER and Golgi in living BY-2 cells ... 80
3.3.5. Mobile Golgi stacks collect ERESs at their rims... 82
3.3.6. Preliminary Characterization of ERESs... 84
4. DISCUSSION ... 87
4.1. Induction and isolation of plant COPII vesicles ... 87
4.2. COPII binding studies... 91
4.3. Visualization of ER exit sites in BY-2 cells ... 94
4.3.1. ERESs in higher plant cells are not organized in transitional ER and do not disappear during mitosis ... 94
4.3.2. Preliminary characterization of higher plant ERESs ... 96
3.3.3. ERESs and the Golgi apparatus: models and data ... 97
3.3.4. ER-to-Golgi transport: vesicles versus tubules... 99
5. REFERENCES... 101
6. APPENDIX ... 124
I. Abbreviation ... 124
II. Frequently used buffers and solutions... 127
III. Sequence alignment ... 129
IV. List of figures... 132
Zusammenfassung
Der Transport zwischen ER und Golgi wird in Hefe und in Säugetierzellen durch COPII- Vesikel vermittelt. Der COPII-Proteinmantel besteht aus der kleinen GTPase Sar1p und den Heterodimerproteinkomplexen Sec23/24 und Sec13/Sec31. Das COPII vermittelte Sorting kommt zustande, wenn die Proteintransporter das ER verlassen. Obwohl die Vorgänge des ER-GA-Transports in Pflanzen ähnlich zu sein scheinen wie in Hefe- und Säugetiersystemen, ist kein Beweis vorhanden, der diese These unterstützt. Mehr noch, es gibt deutliche Hinweise, die nahe legen, dass Unterschiede im Ablauf dieser Prozesse auftreten, wie zum Beispiel das Fehlen des intermediären Kompartiments in Pflanzen, eine große Zahl von Golgiapparaten bewegen sich entlang des Endoplasmatischen Retikulums sowie die Unterschiede in Aufbau und Organisation des Cytoskelets, das an der Interaktion zwischen ER und Golgi beteiligt ist. In dieser Arbeit werden in vitro und in vivo Ergebnisse zum Verständnis der Funktion von COPII-Vesikeln in Pflanzen herangezogen.
In den in vitro vorgenommenen Studien gelang es uns ein Bindungsassay zu entwickeln mit der wir die in vitro Formation von COPII-Vesikeln aufzeigen können. Hierzu wurden
ER-reiche Microsomen, 30% (NH4)2SO4 Zytosol, GMP-PNP und ein ATP regenierendes System
verwandt. Die Vesikelbildung wurde verstärkt, wenn ER-reiche Mikrosomen eines Sec12 Überproduzenten und zusätzliches Sar1p in der Bindungsmixtur verfügbar sind. Potentielle COPII Vesikel wurden von floatierenden Gradienten in der 41 % Saccharosefraktion isoliert, unter dem Elektronenmikroskop stellen sich diese als 50 nm große Vesikel heraus.
Die Fähigkeit des cytosolischen Teils eines pflanzlichen p24 Proteins COPI und COPII- Untereinheiten aus pflanzlichen und tierischen Quellen zu binden, konnte ebenfalls in vitro nachgewiesen werden. Wir fanden ein dihydrophobes Motiv an der -7,-8 Position (relativ zum zytosolischen Carboxyterminus), der für die Bindung der COPII-Untereinheiten sowohl in Arabidobsis als auch im Zytosol von Ratten Leberzellen verantwortlich zu sein scheint. Wie auch immer, anders als im Rattenleberzytosol haben COPI Vesikel aus pflanzlicher Quelle eine stärkere Affinität zum cytosolischen Teil des p24 Proteins als COPII. Nur bei Fehlen des Dilysine-Motivs in der -3,-4 Position (die sehr stark mit dem dihydrophobischen Motiv in der -7,-8 Position interagiert) oder nach dem Entfernen von COPI konnten wir die Bindung von COPII an den p24-Schwanz mit Pflanzenzytosol nachweisen.
Mit dem Hintergrund ERESs in Tabak BY2-Zellen zu visualisieren haben wir zwei verschiedene Methoden entwickelt: a) die direkte Darstellung von endogenen COPII Proteinen (Sar1, Sec13, Sec23) durch Immunfluoreszenzmikroskopie in stabilen Zellinien, die ER- und Golgi lokalisierte GFP-Markerproteine expremieren, und die Darstellung von membrangebundenem Sec13 durch Expression eines Sec13-GFP-Konstrukts in Zellen, die transient ER- und Golgi-lokalisierte RFP-Marker expremieren. In beiden Fällen begrenzen die ERESs erwartungsgemäß die Golgi-Stapel und einige ERESs sind mit Dictyosomen kolokalisiert. Dual-wavelength live cell imaging von ERESs (Sec13-GFP) und Golgi Stapeln (Man1-RFP) zeigt, dass bei Bewegung die Golgi-Stapel ERESs an ihrer Peripherie sammeln. ERES verschwinden nach BFA-Behandlung nicht, stattdessen kommt es zu deutlichen morphologischen Veränderungen im Golgi-Apparat. Die Verhinderung des ER-Exports durch Expression einer Sar1-Mutante, die im GDP-Status angehalten ist, führt zu einem teilweisen Verlust der sichtbaren ERESs.
Summary
Endoplasmic reticulum (ER) to Golgi transport is mediated by COPII vesicles in yeast and mammalian cells. COPII coats consist of the small GTPase Sar1p and the heterodimeric protein complexes Sec23/24 and Sec13/31. COPII-mediated sorting occurs when protein cargo exits the ER. Although the principles of ER-to-GA transport in plant cells are supposed to be similar to those in yeast and mammalian systems, evidence in support of such an assertion is largely circumstantial. Moreover, there is a substantial body of evidence that emphasizes the differences, such as the apparent absence of the intermediate compartment in plants, large numbers of GA stacks moving along the ER and the differences in the organization of the cytoskeleton involved in interrelationships between the ER and GA. In this study, in vitro and in vivo approaches were employed to understand the function of the COPII vesicles in plant cells.
In vitro, we have set up a budding assay which we could monitor the in vitro formation of
COPII vesicle using ER-rich microsomes, 30% (NH4)2SO4 cytosol, GMP-PNP and an ATP
regenerating system. The vesicle budding was enhanced when ER-rich microsomes from a Sec12 overproducer and extra Sar1p are available in the budding mixture. Putative COPII vesicles were isolated from a flotation gradient at 41% sucrose fraction and observed as 50 nm in diameter vesicle under the electron microscope.
The ability of the cytosolic tail of a plant p24 protein to bind COPI and COPII subunits from plant and animal sources in vitro was also examined. We have found that a dihydrophobic motif in the -7,-8 position (relative to the cytosolic carboxy-terminus) is responsible for binding of COPII subunits from both Arabidopsis and rat liver cytosol. However, unlike rat liver cytosol, COPI from plant sources has a stronger affinity for p24 cytosolic tails than COPII. Only in the absence of the dilysine motif in the -3,-4 position (which strongly cooperates with the dihydrophobic motif in the -7,-8 position in binding COPI) or after COPI depletion could we observe COPII binding to the p24 tail with plant cytosol.
In order to visualize ERESs in tobacco BY-2 cells we have employed two different approaches: a) direct visualization of endogenous COPII proteins (Sar1, Sec13, Sec23) in cell lines stably expressing ER- and Golgi-localized GFP-markers by immunoflourescence microscopy, and b) visualization of ER-bound Sec13 by expression of a Sec13-GFP construct in cells transiently expressing ER- and Golgi-localized RFP markers. In both cases ERESs considerably outnumber Golgi stacks, and some ERESs colocalize with Golgi stacks. Dual wavelength live cell imaging of ERESs (Sec13-GFP) and Golgi stacks (Man1-RFP) demonstrates that, as they move, Golgi stacks collect ERESs at their periphery. ERESs do not disappear as a result of BFA treatment, despite considerable morphological changes in the Golgi apparatus. Prevention of ER-export through expression of a Sar1 mutant locked in the GDP state leads to the partial loss of visible ERESs.
1. INTRODUCTION
The eukaryotic secretory pathway sorts and delivers a tremendous variety of proteins to their proper intracellular locations. Delivery proceeds through a series of events including directed membrane translocation, membrane budding and membrane fusion that guide secretory proteins to their ultimate destination. Involved in this process are the normal endomembrane organelles of the endoplasmic reticulum (ER), the Golgi stacks, a cell-delimiting plasma membrane (PM), various endosomal compartments, and a vacuole/lysosome equivalent. Over the past few years, a vast amount of research has illuminated the workings of the secretory system of eukaryotic cells. The bulk of this work has been focused on the yeast Saccharomyces cerevisiae and on mammalian cells. At a superficial level, plants are typical eukaryotes with respect to the operation of the secretory system. In detail, however, important differences emerge in the function and appearance of these organelles in plants. In plants, the ER is at least partially a storage organelle, whereas the Golgi stacks have devoted a major part of their existence to the creation of cell wall precursors. Most visibly, the vacuole has expanded to occupy the majority of the cell volume in mature cells. In addition, the vacuole also has extended beyond the role of a “garbage dump” accorded to the equivalent organelles in other eukaryotes, and has attained major role in the storage of ions, metabolites, and even proteins.
1.1. An overview of the secretory pathway
The endomembrane system is composed of many organelles, each of which must maintain a unique composition of membrane and cargo proteins. Traffic flows in both directions through the secretory system. Anterograde traffic flows from the ER to the PM or vacuole; while retrograde traffic flows counter to the biosynthetic pathway. The transfer of transmembrane proteins and lumenal macromolecules (proteins, glycoproteins, and polysaccharides) between organelles or compartments of the endomembrane system takes place by vesicle transport. Many of the vesicles involved are formed with a protein covering ‘coat’ on their cytoplasmic surface, which later has to be removed to allow for vesicle fusion. These vesicles bud from a donor compartment and then travel to a specific destination where they fuse with the target compartment. Clearly, some manner of regulation is required to prevent mistargeting of vesicles to an incorrect compartment.
Intracellular transport between early compartments of the secretory pathway relies on a series of protein-sorting events that are accomplished by coat protein complexes
(COPs). In general, activated small guanine triphosphatases (GTPases) recruit coat proteins to specific membrane export sites, thereby linking COPs to export cargos. As coat proteins polymerize, vesicles are formed and bud-off from membrane-bound organelles. Transport between the endoplasmic reticulum (ER) and Golgi, traffic is bidirectional, ensuring that proteins required to form and fuse vesicles with organelles are recycled as secretory cargo advances. COPII vesicles bud from the ER for anterograde transport, whereas the COPI coat appears to be responsible for retrograde transport of recycled proteins from Golgi and pre-Golgi compartments back to the ER (Figure 1.1).
Fig. 1.1 Intracellular Transport Pathways
The scheme depicts the compartments of the secretory, lysosomal/vacuolar, and endocytic pathways. Transport steps are indicated by arrows. Colors indicate the known or presumed locations of COPII (blue), COPI (red), and clathrin (orange). Clathrin coats are heterogeneous and contain different adaptor and accessory proteins at different membranes. Only the function of COPII in ER export and of plasma membraneassociated clathrin in endocytosis are known with certainty. Less well understood are the exact functions of COPI at the ERGIC and Golgi complex and of clathrin at the TGN, early endosomes, and immature secretory granules. The pathway of transport through the Golgi stack is still being investigated but is generally believed to involve a combination of COPI-mediated vesicular transport and cisternal maturation (Pelham and Rothman, 2000). Additional coats or coat-like complexes exist but are not represented in this figure. (From Bonifacino and Glick 2004)
1.2. Secretion-related Organelles and Compartments in Plant
Cells
1.2.1. Endoplasmic Reticulum (ER)
Proteins intended for the endomembrane system typically contain N-terminal signal peptides, or have analogous transmembrane domains at other places within the protein that engage the ER-translocation machinery in a similar way (Martoglio and
Dobberstein, 1998). The overall mechanism and the proteinaceous machinery is remarkably conserved across eukaryotes. Most eukaryotes are capable of both co- and post-translational translocation, and often similar proteins may use either method depending on the species in which it is expressed (Kalies and Hartmann, 1998). Regardless, signal peptides can be successfully exchanged between different organisms (Gierasch, 1989), indicating the overall conservation of the process.
Once engaged by the translocation machinery, proteins that lack membrane-spanning domains are released into the lumen of the ER, whereas the membrane proteins are threaded into the membrane (often multiple times) depending upon the folding information within the peptide sequence of the membrane protein. In either case, chaperones and other factors assist in the folding, disulfide bond formation, core-glycosylation, and oligomerization of the proteins. During this process, misfolded proteins are selectively retained by a quality control process which either completes the folding and releases the protein, or marks it for destruction (Vitale and Denecke, 1999).
The ER is comprised of a three-dimensional network of continuous tubules and sheets that underlies the plasma membrane, courses through the cytoplasm, and links up with the nuclear envelope. It is the most versatile and adaptable organelle of eukaryotic cells. Its principal functions include the synthesis, processing and sorting of proteins, glycoproteins and lipids, ER associated protein degradation, as well as the regulation of cytosolic calcium levels. The classical literature distinguishes three ER subcompartments: rough ER, smooth ER, and the nuclear envelope. However, it has been suggested that the ER has many distinct domains based upon metabolic, morphological, and other criteria (Figure 1.2. Staehelin, 1997). The tER is the domain where secretory cargo proteins become concentrated for packaging into COP-II transport vesicles which will carry them to the Golgi stacks. tER has been best studied in mammals and fission yeast, where clear morphological and cytochemical evidence for tER “exit sites” is found (Hammond and Glick, 2000, Rossanese et al., 1999). Similar sites have seldom been reported in plant cells (Staehelin, 1997). The tER-type domain may not be strictly necessary, as baker’s yeast does not appear to produce specific tER sites, and instead, it is believed that cargo may exit from any point in this type of yeast (Rossanese et al., 1999).
Packaging of cargo in the tER still remains somewhat controversial. While some of the proteinaceous components of the vesicle forming machinery have been characterized, it is still unclear whether there is a selective packaging or simple “bulk-flow” of lumenal contents towards the Golgi. Many so called ER-to-Golgi “cargo
receptors” have been identified in yeast and mammals suggesting specific packaging (Lavoie et al., 1999, Denzel et al., 2000,etc.). Whether or not a bulk-flow or specific mechanism exist in plant cells has recently been investigated (Phillipson et al., 2001; Törmäkangas et al., 2001), and further work will be required to implicate one over the other. Despite some level of specificity in packaging, ER-residents are often reported to escape as far as the trans-Golgi network, and an efficient Golgi-to-ER recycling system (KDEL-receptor) has been found for their retrieval (Vitale and Denecke, 1999). Thus, it is likely that some selective process is functioning for particular proteins, whereas some nonselective sampling of the lumenal contents also occurs. Packaging of membrane proteins is likely to be more selective and is probably dependant on peptide signals within the membrane spanning domain.
Fig. 1.2. Schematic diagram of a plant cell depicting the sixteen types of ER domains.
PM, plasma membrane; TV, transport vesicle; SV, secretory vesicle; TGN, trans Golgi network.(From L.A.Staehelin，1997)
The Golgi is not the only destination for cargo exiting from the ER. Many plant species create unique protein bodies of seed storage proteins by selective distention of an ER subdomain (Herman and Larkins, 1999). Evidence for a direct (non-Protein Body) pathway from the ER to the vacuole has also been reported (Jiang and Rogers, 1998), although mechanistically, little else is known about this type of trafficking. It
has recently become clear that peroxisomes may also be formed (at least partially) by budding from a sub-domain of the ER (Mullen et al., 2001).
1.2.2. Golgi Apparatus (GA)
The plant Golgi apparatus shares many features with its animal counterpart, but also has unique characteristics. The most important difference concerns its structure. Whereas in animal cells the Golgi apparatus occupies a rather stationary perinuclear position, in plant cells the Golgi is divided into individual Golgi stacks, which are generally considered to functionally independent (Staehelin and Moore, 1995). Each stack is typically formed of 5-10 individual cisternae. The number of Golgi stacks per cell and the number cisternae per stack vary with the species and cell type, but also reflect the physiological conditions, the developmental stage and the functional requirements of a cell (Staehelin and Moore, 1995; Andreeva et al., 1998). Despite these variations, each individual Golgi stack can be described as a polarized structure in term of cisternal morphology and with enzymatic activities changing gradually from the ER-adjacent cis-face to the trans-face (Fitchette al., 1999). Proteins destined for secretion enter the Golgi at the cis-face and subsequently move towards the trans-face where the majority of proteins exit the stack. Although no matrix proteins surrounding the plant Golgi stack have yet been identified, the existence of a matrix has been predicted from the appearance of a clear zone, excluding ribosomes, around each Golgi on micrographs from ultra-rapidly frozen root cells (Staehelin et al., 1990). The Golgi matrix has been suggested to play an important role in the maintenance of stack organization against the shearing forces during cytoplasmic streaming (Staehelin and Moore, 1995)
Confocal microscopy of Golgi-targeted proteins or peptides fused to green fluorescent protein (GFP) has revealed that individual stacks are highly mobile within the plant cell, moving over the ER on an actin-network (Boevink et al., 1998; Nebenführ et al., 1999; Brandizzi et al., 2002). This has resulted in them being christened ‘stacks on tracks’ (Boevink et al., 1998) or ‘mobile factories’ (Nebenführ and Staehelin, 2001). The fact that the plant Golgi apparatus is divided into highly mobile biosynthetic subunits certainly poses major problems when trying to elucidate mechanisms for controlled protein import into and targeted product export out of the stacks.
How the cargo passes through the cisternae is still a matter of some controversy (Pelham and Rothman, 2000). Some evidence suggests that the cisternae are static organelles, and that all traffic between the stacks is vesicle-mediated by COP-I coated vesicles. Other evidence indicates that the stacks may simply be transient structures
that are formed new from ER-derived vesicles at the cis-side, then sequentially mature into medial- and trans-cisternae (cisternal maturation model). New cis-stacks form behind the maturing cisternae creates an “assembly-line”-like formation of stacks. The Golgi is the first cisterna encountered following the ER. Within the cis-cisterna, modifications to the core N-glycosylation of proteins begin. Enzymes such as α-mannosidase remove the terminal mannose residues, creating substrates for other glycosyltransferases which act in the later cisterna to produce the unique glycosylation patterns found on many proteins. This enzyme, α –mannosidase I, has been fused to green fluorescent protein (GFP) by the Stahaelin group, and has been used as an in vivo marker for the Golgi stacks, revealing some interesting behavior in dividing cells (Nebenführ et al., 2000 ).
The cis-cisternae is also the site of recapture of escaped ER residents. The KDEL-receptor is a transmembrane protein whose lumenal domain specifically recognizes a C-terminal K/HDEL motif found at the C-terminus of ER-resident soluble proteins (Pelham, 1996). Upon binding of proteins with these signals, the KDEL-receptor recruits the COP-I machinery and mediates the return of these proteins to the ER. Recent work has indicated that the plant KDEL receptor appears to exceptionally efficient, and rarely allows any ER residents to escape past cis-cisterna (Phillipson et al., 2001). In contrast, experiments in mammalian cells have indicated that ER residents may sometimes be allowed to pass as far as the TGN before recapture. Since the number of cisternae in a Golgi apparatus can vary significantly, definition of a “medial” stack is equally variable. Enzymatically, these stacks can often have activities that differ from that of the earlier and later cisternae. Certainly, the spectrum of glycosyltransferases must be somehow distinct, but how this distinction is setup and maintained remains unclear. Some evidence suggests that synthesis of some cell wall glycans is initiated in the medial stacks, whereas the trans-cisterna is the typical site of xyloglucan assembly (Dupree and Sherrier, 1998). Some of the molecular details of the glycosyltransferasese involved in these processes have begun to be worked out, though much remains to be done (Steinkellner and Strasser 2003).
The ELP/BP-80 class of vacuolar sorting receptors (VSR) are found concentrated in the trans-cisternae and the TGN (Paris et al., 1997; Sanderfoot et al., 1998). These proteins specifically recognize one particular class of vacuolar sorting signal (NTPP or sequence-specific VSS), leading to a concentration of these proteins to specific domains of the TGN (Kirsch et al., 1994 ; Cao et al., 2000 ; Ahmed et al., 2000). The VSR then recruits a clathrin-type coat to the membrane, and directs the packaging of the cargo into CCVs. These CCVs travel onto the prevacuolar compartment. Storage proteins are not recognized by the ELP/BP-80 class. In certain cell types, these types
of proteins are packaged into “Dense Vesicles” (Hara-Nishimura et al., 1998; Hinz et al., 1999; Hillmer et al., 2001). The molecular details of this pathway are not yet clear, but a condensation-sorting mechamism as known for regulated secretory proteins in mammals has been suggested(Robinson and Hinz 1999).
Another aspect of Golgi mediated targeting occurs during cytokinesis. Plants have a unique method of cell division whereby a novel membrane structure (the cell plate) is synthesized by Golgi-derived vesicles at the point of cytokinesis. The cell plate is a site of intense vesicle fusion and formation, and eventually, a new plasma membrane is formed by the fusion of the cell plate with the maternal plasma membranes. Plant cytokinesis is a distinct membrane trafficking process in at least three ways. First, this process is confined to the mitotic phase of the cell cycle whereas other trafficking processes occur during interphase when the cell is growing in size. Second, unlike vesicles destined to fuse with target membranes in the non-dividing cell, cytokinetic vesicles initially fuse with one another to form a new membrane compartment, the cell plate, which becomes the target membrane for fusion of later arriving vesicles. Third, vesicle trafficking during is assisted by a plant-specific cytoskeletal array, the phragmoplast, that forms only in late anaphase and disassembles upon the completion of cytokinesis (Jürgens and Pacher 2003).
1.2.3. Prevacuolar Compartment (PVC)
Prevacuolar compartments (PVCs) are defined as organelles that receive cargo from transport vesicles and subsequently deliver that cargo to the vacuole by fusion with the tonoplast (Bethke and Jones 2000). Based on precedents from mammalian and yeast systems (Lemmon and Traub 2000), they are intermediate organelles on the pathways to vacuoles from both Golgi and from endocytosis of the plasma membrane. In mammalian cells, they have been identified as the prelysosomal compartment (PLC). It is a 0.5-1 µm diameter organelle, which typically has internal 60-80 nm vesicles (Dunn et al., 1986, van Deurs et al., 1993).
There is increasing evidence suggesting that PVCs exist in plant cells and they play a similar role in protein trafficking in the plant secretory pathway (Robinson and Hinz 1999, Bethke and Jones 2000). Identification of these PVCs will enable functional definition of their roles in the complex plant vacuolar system and the vesicular pathways leading to multiple vacuoles (Jiang and Rogers 1999, Vitale and Raikhel 1999, Robinson et al., 2000).
Compared to the yeast and mammalian cell, plant cells contain two functionally distinct vacuoles: the protein storage vacuole (PSV) and lytic vacuole (Hoh et al., 1995, Paris et al., 1996 Vitale and Raikhel 1999). Putative PVCs for the PSVs have been identified and characterized. In developing pea seeds, multivesicular bodies (MVBs) rangeing in size from 0.5 to several µm containing storage proteins have been postulated as PVCs for PSVs (Robinson et al., 1998). Although, there are some evidences support the existence of lytic PVC (Li et al., 2002). Whether or not there are two functional distinct types of PVCs in plant cells is not clear. Several proteins markers like BP-80 (Kirsch et al., 1994), AtPep12 (Conceicao et al., 1997) and AtELP (Sanderfoot et al., 1998) specific for the PVC have been reported. However, due to the complexity of the plant late endosome system, there are multiple pathways using distinct transport vesicles for transporting proteins to the PSV and lytic vacuole in plant cells (Hara-Nishimura et al., 1998, Hinz et al., 1999, Jiang and Rogers 1999, Vitale and Raikhel 1999). The PVC is still not well defined.
Receptor-mediated endocytosis is a well-documented feature of mammalian cells (Trowbridge et al., 1994, Mukherjee et al., 1997), and has also been established for yeast cells (Geli and Riezman, 1998). In both case, the biosynthetic (TGN-PLC/PVC-lysosome/vacuole) and endocytotic pathways to the vacuole converage at the PLC/PVC. However, receptor-ligand complexes which are internalized at the plasma membrane via CCV are not delivered directly to the PLC/PVC since they are on the way to the degradatic compartment. At certain point the receptor must be recycled to the plasma membrane. This occurs at the compartment called early endosome, in contrast to the PLC/PVC, which is termed the late endosome.
Three possibilities exist for the arrival of the contents and the membrane of the PLC/PVC into the lysosome/vacuole: maturation and gradual transformation of the PLC/PVC into lysosome/vacuole (Murphy, 1991); fusion of the PLC/PVC with a pre-existing lysosome/vacuole (Griffiths and Gruenberg, 1991); and by vesicle transport.
1.2.4. Vacuoles
In a plant cell, the most obvious organelle is the large central vacuole (Marty, 1999). Initially thought of as a large empty space, the central vacuole often occupies as much as 90% of the cell volume in a mature cell (Cutler et al., 2000). The vacuole is also the equivalent of the lysosome of mammals, in that it contains the various hydrolyases capable of degrading and recycling proteins, lipids, carbohydrates. In concert with the plasma membrane, the vacuole also has the essential role of maintaining turgor
pressure in the plant cell. This turgor is essential for homeostasis and also for growth and development in plant. Recently, this has been underlined is a paper by Rojo et al. (2001) with the vacuoleless mutant of Arabidopsis. Plants where the Vacuoleless gene is disrupted by a T-DNA insertion were found to completely lack a central vacuole, and die as embryos.
Recent research has rediscovered an old observation in plant cytology: a single plant cell can have more than a single type of vacuole (Hoh et al., 1995; Paris et al., 1996; Di Sansebastiano et al., 1998; Swanson et al., 1998). It has long been observed that in the storage cells of embryo-derived tissue a new organelle develops that contains storage proteins. This structure was called either a protein body (PB) or a protein storage vacuole (PSV), depending on the species of plant. In garden pea (Pisum
satium), the storage proteins destined for the PSV begin to condense in the early
stacks of the Golgi and form “Dense vesicles” (Hohl et al., 1996; Hinz et al., 1999; Hillmer et al., 2001). These DVs travel from stack-to-stack at the distal ends of the cisterna – or alternately, are carried along the maturing cisterna as it travels forward. Either way, the DV expands as additional cargo is packaged within, until finally budding free from the TGN. The DV is subsequently trafficked to a structure similar to a MVB, before eventually fusing with the PSV (Robinson et al., 2000).
In other species (the cucurbits), storage proteins are instead packaged into a DV-like organelle (called a PAC-precusor accumulating) at the ER (Hara-Nishimura et al., 1998). The ER distends until a PAC buds free into the cytoplasm. A single PAC may fuse with other PACs, receive additional cargo delivered from the Golgi, eventually form an MVB-like organelle, and then fuse with a PSV.
1.3. Vesicles mediating cargo transport in the early secretory
pathway
Intracellular protein transport in eukaryotic cells is mediated by small transport vesicles that are defined by their coat proteins: COPII-coated vesicles allow exit from the endoplasmic reticulum (ER), COPI vesicles carry proteins within the early secretory pathway (i.e. the ER and Golgi apparatus) and clathrin-coated vesicles mediate transport from the trans-Golgi network (TGN) and endocytic transport from the plasma membrane (Schekman and Orci, 1996; Rothman and Wieland, 1996; Schmid, 1997; Barlowe, 1998). Vesicular transport intermediates not only perform delivery of cargo to various destinations: at the same time they regulate the steady state of the endomembrane system of a eukaryotic cell (Wieland and Harter 1999).
Formation of a transport vesicle is not a spontaneous event. A mechanism for cargo selection and concentration is required to decide what belongs in which vesicle. Secondly, some signal must be sent from the donor membrane into the cytoplasm (where a coat must form) to indicate a site for vesicle formation. Most importantly, some method of membrane deformation and scission from the donor compartment is needed to actually produce the vesicle. This is the job of a large collection of proteins that are referred to as coat proteins. Each type of coat is distinct, though some are related, and a particular coat is responsible for vesicle formation at a particular type of organelle. An overall similarity in the coating mechanism is found, even though the protein content of the coats may vary. The first step requires recruitment of the cargo to a site on the donor membrane, the details of such a step is still a matter of debate. However, this step most likely involves the cytoplasmic tails of integral membrane proteins. These proteins may themselves be cargo, or may serve as cargo receptors. Coincident with this step is the GTP-cycle of a coat-GTPase which helps to coordinate the coating process. The coat-GTPases typically exist as a soluble GDP-bound state while inactive. A GTP-exchange factor (GEF) localized to the coating site serves to recruit the coat-GTPase to the donor membrane and then triggers a nucleotide exchange, such that the coat-GTPase assumes its active GTP-bound state. The GEF for the Sar1p-type G-protein is a member of the Sec12-family, a group of ER-integral membrane proteins found in all eukaryotes. The GEF for the ARF-type G-protein vary somewhat, since the ARF-type coat-GTPase is involved in many different coating steps (Kirchhausen, 2000).
Subsequently, a GTPase Activating Protein (GAP) triggers the intrinsic GTPase activity of the coat-GTPase. This step occurs either directly after vesicle formation (COP-II) (Antonny et al., 2001), or can be triggered by some other event (COP-I or clathrin) (Tanigawa et al 1993; Cukierman et al., 1995). The GDP-bound G-protein now triggers uncoating of the vesicle, releasing the coatomers back to the cytoplasm for future coating steps. The uncoated vesicle is now free to travel to its target membrane where the SNARE-mediated fusion process occurs.
1.3.1. COPI Vesicles
COP I-coated vesicles were first identified from an intra-Golgi transport assay (Balch et. al., 1984). In this assay their formation was observed when Golgi membranes were incubated in the presence of cytosol, fatty acyl-coenzyme A, and nucleotides (Balch et. al., 1984; Pfanner et al., 1989). They accumulated under conditions known to block protein transport such as the addition of non-hydrolysable GTP analogues (Melancon et al., 1987). This led to the hypothesis that they were the vesicles mediating forward
transport through the Golgi complex. Subsequently these vesicles, now known as COP I vesicles, were purified to homogeneity (Malhotra et al., 1989) and the cytosolic components forming their coat characterized (Serafini et al., 1991; Water et al., 1991). The complex these components form in the cytosol was named the ‘coatomer’ and consisted of 7 polypeptides plus the small GTPbinding protein ARF1 that is not part of the coatomer itself. There are two conformations of coatomer: that of the soluble coatomer; and that of coatomer in COPI-coated vesicles or aggregates. Coatomer aggregation in the presence of tetrameric p23-CT thus appears to be related to COPI coat polymerization on native membranes, which suggests that the interaction of p24 proteins with coatomer is a critical trigger of COPI coat assembly (Harter and Wieland, 1998; Reinhard et al., 1999).
COPI vesicles instead appear to be involved in both biosynthetic (anterograde) and retrograde transport within the Golgi complex (Orci et al., 1997), as well as mediating the recycling of proteins from the Golgi to the ER (Cosson and Letourneur, 1994; Letourneur et al., 1994; Sönnichsen et al., 1996).COP-I coats are typically associated with Golgi trafficking, although they may also be involved in forming coated vesicles at the ER and endosomal compartments (Kirchausen, 2000). The COP-I coatomer is made up of seven components (α /Rer1p, β/Sec26p, β'/Sec27p, γ/Sec21p, δ/Ret2p, ε/Sec28p, and ζ/Ret3p; listed in the mammalian/yeast nomenclature) that are conserved across eukaryotes (Table 1.1). As with COP-II coatomers, the COP-I-coatomers are encoded by multiple genes (except for γ and δ) in Arabidopsis. The GTPase that drives COP-I coat formation is from the ARF-family, a large family of small GTPases (at least 18 in Arabidopsis) that also function in other coating processes. The COP-I coatomer also plays a role in cargo selection, with various members of the complex interacting specifically with motifs present in cytoplasmic domains of cargo proteins. COP-I coated vesicles seem to be involved in both anterograde and retrograde trafficking between the Golgi stacks – though a role in anterograde trafficking would be unnecessary in a cisternal maturation mode of Golgi functioning.
The initial event in the COPI pathway that leads to recruitment of the coat requires the association of the GTPase ARF1(ADP-ribosylation factor 1) in its active form to the membrane.The ARF protein family has many members and targeting of ARF1 to the correct membrane involves specific association with its appropriate GEF. Several GEFs have been identified that stimulate the guanine nucleotide exchange rate of specific ARFs, and each possesses a 200-amino acid segment referred to as the ‘Sec7 domain’ (Chardin et al., 1996; Chardin et al., 1999). Interestingly, the ARF1–Sec7 domain complex resembles more closely the GTP and not GDP conformation, suggesting that some form of ARF1 activation precedes interaction with the Sec7 domain.
Recent progress has also been made in understanding the molecular detail of brefeldin A (BFA) effects on COPI coat assembly. It had been shown that BFA addition to certain cell types leads to a disassembly of the Golgi complex and redistribution into the ER (Lippincott-Schwartz et al., 1989; Klausner et al., 1992). Specifically, BFA inhibits the rate of guanine nucleotide exchange exhibited by ARF1, and interferes with ARF1 function in recruiting the COPI coat (Donaldson et al., 1992; Helms and Rothman 1992). In essence, BFA acts by sequestering specific Sec7 domain-containing proteins, which in turn prevent the activation of ARF1 and ultimately blocks COPI coat formation.
ARF-specific GAPs have been identified that accelerate the rate of GTP hydrolysis. A founding member of this class of proteins contains a zinc finger motif that is essential for ARF GAP activity and stimulates the rate of GTP hydrolysis (Cukierman et al., 1995).
COPI-coated vesicles efficiently capture proteins carrying in their cytoplasmic carboxyterminal domain sorting signals of the form KKXX (the dilysine motif) or KXKXX (X is any amino acid). The KDEL receptor, a multiple-spanning membrane protein that binds and retrieves lumenal proteins containing the KDEL carboxy-terminal sequence, is also transported along this pathway. The γ subunit seems to be the component responsible for cargo recognition because it recognizes the KKXX and KXKXX motifs, but it is not known whether it also recognizes the KDEL receptor.Members of the p24 protein family also interact with COPI coatomers in addition to COPII and might facilitate the recruitment of COPI coatomers to Golgi membranes (Dominguez et al., 1998). p24 proteins not only bind to coatomer and alter its conformation but are also abundant residents of the intermediate compartment (IC) and the Golgi (Sohn et al., 1996; Rojo et al., 1997; Dominguez et al., 1998),
which are the intracellular sites of COPI vesicle biogenesis (Griffiths et al., 1995; Orci et al., 1997).
Therefore, it has been proposed that the minimal machinery for COPI vesicle formation consists of p23/p24, ARF-GTP and coatomer (Figure 1.3; Nickel et al., 2002; Aniento et al., 2003).
Fig. 1.3 The core machinery of COPI recruitment to membranes, coat polymerization,
vesicular budding and uncoating.
Recruitment of coat proteins is initiated by ARF-GDP binding to p23 (1). Upon nucleotide exchange, ARF-GTP dissociates from p23 resulting in its stable association with the membrane (2). Multiple cycles of GTP hydrolysis and GDP to GTP exchange are likely to occur, possibly causing rearrangements of p23/p24 oligomers (3). The products of these processes are ARF-GTP and presumably a p23/p24 heterooligomer, which triggers coatomer binding and coat polymerization (4). Following budding (5), the catalytic domain of ARF-GAP is sufficient to trigger uncoating (6). Active components are shown in green; inactive components are shown in red. (From Nickel et al., 2002)
1.3.2. COPII Vesicles
The second class of vesicle is the COP II vesicle identified throught a combination of genetical and biochemical approaches in the yeast Saccharomyces cerevisiae (Antonny and Schekman, 2001). These vesicles mediate the transport of a subset of secretory proteins from the ER to the Golgi complex.
The COP-II coat is conserved in all eukaryotes and is involved in transport from the ER. In most organisms, ER-exit sites are discrete points along the ER membrane (Bannykh and Balch, 1997; Rossanese et al., 1999), although in some organisms (like
2001). The exit sites are points at which the COP-II machinery is concentrated (Hammond and Glick 2000), and also where anterograde cargo is collected. How cargo becomes concentrated remains unclear, with some arguing for a “bulk-flow” model, whereas other invoke specific cargo receptors. In a bulk-flow model, all cargo not specifically retained in the ER is subject to COP-II-directed (Herrmann et al., 1999) transport to the Golgi. On the other hand, some propose that a large class of conserved proteins are involved in clustering cargo for packaging at exit sites. Evidence for both models has accumulated, and it is possible that both occur depending on the cell-type or developmental state or particular cargo molecule.
1.3.2.1. COPII coat components
The COP-II coat is made of 5 proteins (Antonny and Schekman, 2001). Having been first described in yeast, the components are typically referred to according to the yeast nomenclature. Once activated by the Sec12-GEF, Sar1p-GTP recruits two coatomer protein complexes from the cytosol: the Sec13/31p and the Sec23/24p complexes (Table 1.2). The Arabidopsis coatomer subunits are each encoded by multiple genes, though whether this indicates redundancy has yet to be investigated. The coatomer complexes then assemble into higher order structures which drive membrane deformation and eventual membrane budding. The COP-II coated vesicles are quite unstable as the GAP for Sar1p turns out to be a component of the coat itself (Sec23p), thus the vesicles are uncoated soon after budding. These vesicle quickly fuse with each other into larger vesicles which either fuse with the cis-Golgi (or form a de novo cis-Golgi stack in cisternal maturation models), or with a compartment called the ER-Golgi Intermediate complex (ERGIC) found in most mammals. Upon uncoating, Sar1p and the coatomers are released into the cytoplasm for further rounds of vesicle budding.
Table 1.2. Coat proteins of COPII vesicles (*Arabidopsis isoforms)
Sar1p
The Sar1 GTPase is an essential component of COPII vesicle coats involved in export of cargo from the ER. The GTPase activity of Sar1 function as a molecular switch to control protein-protein and protein-lipid interactions that direct vesicle budding from the ER (Springer et al., 1999). Activation from the guanosine 5′-diphosphate (GDP) to the GTP-bound form of Sar1 involves the membrane-associated guanine nucleotide exchange factor (GEF) Sec12, first identified in yeast (Nakano et al., 1988; Barlowe et al., 1993). A homologue of yeast Sec12 has also identified in plants (Bar-Peled and Raikhel, 1997) and in mammalian cells (Weissman et al., 2001). During or after fission of vesicle from ER, GTP hydrolysis is stimulated by the recruited Sec23/24 GTPase-activating protein (GAP) complex to promote coat disassembly and subsequent delivery of cargo to the Golgi complex (Barlowe et al., 1994).
The GDP-binding site includes residues found in all of the highly conserved binding motifs diagnostic of GTPases.Thr39 is found in the conserved GxxxxGKT39 motif. It is involved in the co-ordination of a Mg2+ ion, which is also co-ordinated to oxygen atoms of the ß- and γ-phosphates of GTP (Pai et al., 1990) and is essential for Sar1 function. In particular, the Sar1 dominant negative mutant Sar1 [T39N] is a potent inhibitor of COPII vesicle formation (Kuge et al., 1994; Aridor and Balch, 1996; Rowe et al., 1996). Sar1 [T39N] inhibits wild-type Sar1 function in vivo and in vitro by interfering with its interaction with mammalian Sec12 (mSec12), the ER-associated GEF that is required for Sar1 activation and COPII vesicle formation (Weissman et al., 2001).
A second mutation, Sar1 [H77L], abolishes the His residue that presumably coordinates the water molecule participating in GTP hydrolysis, and blocks the GTPase activity of Sar1 leading to its stabilization in the GTP-bound activated form. The Sar1-[H77L] mutant supports vesicle budding but displays no overall ER-to-Golgi transport in cell –free assays. It is insensitive to the GAP function of the sec23/24 complex (Saito et al., 1998).
Sec23/24 complex
The Sec23p/24p complex is a heterodimer. Sec23p/24p elutes on gel filtration chromatography as a single species corresponding to a globular protein of about 200 kDa. The masses of Sec23p and Sec24p deduced from their cDNAs are 85 and 104 kDa, and the molar ratio of Sec23p to Sec24p in the complex is 1:1, as assessed by a
combination of SDS/PAGE and quantitative amino acid analysis. The Sec23/24 complex has a bone-like outline, the outside dimensions of which are about 11x17 nm (Lederkremer et al., 2001).
The Sec23p–Sec24p complex is probably the component responsible for cargo recognition (Springer and Schekman 1998; Peng et al., 2000) but the sorting signals recognized by the complex remain to be identified. Members of the p24 family of transmembrane proteins bind to Sec23p through a cytosolic diphenylalanine motif. As these proteins are required for efficient ER-to-Golgi traffic of some cargo proteins (Schimmoller et al., 1995), it is thought that they might serve as cargo adaptors (Kaiser 2000). In addition to recruiting the Sec23p–Sec24p complex, the GTP-bound form of Sar1p activates Sec23p to bind SNARE proteins involved in the specificity of targeting and in the fusion reaction of vesicles with acceptor membranes (Springer and Schekman 1998). ER membranes with Sec23p–Sec24p and Sar1p can then recruit Sec13p–Sec31p. The complex is likely to act as a scaffold to drive membrane deformation and to complete vesicle budding. Sec23p also acts as a GTPase-activating protein (GAP) for Sar1p. It is thought that, after GTP hydrolysis, Sar1p– GDP is released, leading to uncoating before fusion of the vesicle to the target membrane and formation of a new coated vesicle.
Sec13/31 complex
The Sec13p/31p complex is a heterotetramer. During gel filtration chromatography, Sec13p/31p eluted as a single monodisperse species corresponding to a globular protein of about 700 kDa in mass. Because of the marked disparity with the added molecular masses of its Sec13p and Sec31p subunits (33 and 140 kDa, respectively), it has been proposed that the Sec13p/31p complex might be a heterodimer of elongated shape (Salama et al., 1997), although the possibility that the complex could contain more subunits was not explored. Sec13p/31p is a relatively asymmetric heterotetramer. The simplest organization that is consistent with the elongated shape of Sec13p/31p is a side-by-side arrangement of its subunits, such that the globular domain at one end of the complex corresponds to two Sec13p subunits, and the globular domain at the opposite end contains the carboxyl terminal part of Sec31p (Lederkremer et al., 2001).
1.3.2.2. COPII formation and disassembly
Vesicular transport can be reconstituted by using three cytosolic components containing five proteins: the small GTPase Sar1p, the Sec23p/24p complex, and the
Sec13p/Sec31p complex (Salama et al., 1993). These proteins will support a cargo-carrying budding reaction from isolated ER membranes. Sar1p, a GTP-binding protein, initiates coat formation (Matsuoka et al., 1998). The GDP-bound form of Sar1p is normally cytosolic. It is recruited to the ER membrane by interaction with Sec12p, an ER-bound membrane protein that serves as its guanine exchange factor (Barlowe et al., 1993). Sar1p-GTP then recruits cytosolic Sec23p/24p complex, most likely through its interaction with Sec23p (Yoshihisa et al., 1993).
In addition to recruiting Sec23p/24p, the GTP-bound form of Sar1p stabilizes Sec23p and binds to certain ER and Golgi SNARE proteins involved in the specificity of targeting and in the fusion reaction of vesicles with acceptor membranes (Springer and Schekman 1998). The interaction of Sar1p-GTP with Sec23p also facilitates the association of the Sec23p/24p complex with cargo proteins (Kuehn et al., 1998); Sec24p is probably the component responsible for cargo recognition (Shimoni et al., 2000). ER membranes with Sec23p/24p and Sar1p can then recruit Sec13p/31p, a complex that is likely to act as a scaffold, like clathrin, to effect membrane deformation and vesicle budding. Completing the cycle, Sec23p acts as a GTPase activating protein for Sar1p (Yoshihisa et al., 1993). It is thought that on GTP hydrolysis; Sar1p-GDP is released, leading to uncoating before fusion of the vesicle to the target membrane and recycling of COPII components.
Fig. 1.4. Assembly of COPII
Cytosolic Sar1p-GDP is converted to membrane bound Sar1p-GTP by the transmembrane protein Sec12p. Sar1p-GTP recruits the Sec23p/Sec24p subcomplex by binding to Sec23p, forming the “pre-budding complex”. Transmembrane cargo proteins gather at the assembling coat by binding to Sec24p. The Sec13p/Sec31p subcomplex polymerizes onto Sec23p/Sec24p and crosslinks the pre-budding complexes. Cargo proteins are further concentrated. The depictions of Sar1p, Sec23p, and Sec24p are surface representations from the crystal structures of these proteins (Bi et al., 2002). The Sec13p/Sec31p complex is represented as an elongated, five-globular domain structure based on electron microscopy (Lederkremer et al., 2001). Sec16p and Sed4p also participate in the assembly of COPII, but are not represented here because their roles are less well understood. See text for additional details. (From Bonifacino and Glick 2004)
At present, it is unclear what marks ER exit sites for COPII recruitment. A candidate for this role is Sec16p, a large peripheral ER membrane protein (Espenshade et al., 1995). Sec16p interacts with Sec23p, Sec24p, and Sec31p via different domains (Espenshade et al., 1995; Shaywitz et al., 1997) and may serve as scaffold for the nucleation or stabilization of the assembling coat (Supek et al., 2002). It is likely that Sec16p acts in conjunction with the transmembrane protein Sec12p to recruit GTP bound Sar1p to the ER membrane. Sar1p-GTP associates with the lipid bilayer through a hydrophobic amino-terminal extension and recruits its effector, the Sec23p/Sec24p subcomplex, through interactions with two “switch” regions characteristic of Ras superfamily proteins (Huang et al., 2001; Bi et al., 2002). The initiation of COPII assembly thus involves both independent and GTP-dependent reactions that cooperate to deposit the coat at ER exit sites.
Sar1p-GTP together with Sec23p/Sec24p constitutes the so-called “pre-budding complex,” which has recently been analyzed by electron microscopy (Lederkremer et al., 2001; Matsuoka et al., 2001) and X-ray crystallography (Bi et al., 2002). This complex has the appearance of a bow tie with one side corresponding to Sec23p and the other to Sec24p (Bi et al., 2002). Sec23p makes direct contact with Sar1p-GTP (Bi et al., 2002), while Sec24p participates in cargo recognition. Once assembled onto membranes, the pre-budding complex recruits the Sec13p/Sec31p subcomplex, which consists of two Sec13p and two Sec31p subunits (Lederkremer et al., 2001). Sec13p/Sec31p appears by electron microscopy as a flexible, elongated structure that polymerizes to form a meshlike scaffold (Lederkremer et al., 2001; Matsuoka et al., 2001).
Sec23p stimulates the GTP hydrolysis activity of Sar1p (Yoshihisa et al., 1993) by contributing an “arginine finger” that pokes into the GTP binding site and aids catalysis (Bi et al., 2002). This activity of Sec23p as a GTPase-activating protein (GAP) is augmented approximately ten-fold by addition of Sec13p/Sec31p (Antonny et al., 2001). A paradoxical implication of this mechanism is that COPII coat assembly should trigger disassembly by promoting GTP hydrolysis. How can the COPII coat polymerize to cover a forming vesicle if the basic unit of the polymer is unstable? A possible explanation is that the kinetics of GTP hydrolysis might be slower than the kinetics of vesicle budding, in which case there would be time for a vesicle to form before the coat fell apart. Alternatively, GTP hydrolysis might cause Sar1p to be released from the coat while the other subunits remained assembled on the membrane. The polymeric nature of the coat could provide kinetic stability in the absence of Sar1p-GTP. In addition, the cytosolic domains of transmembrane cargo
proteins could act as secondary membrane tethers or could modulate the GAP activity of Sec23p. Any of these alternative explanations would imply that Sar1p-GTP is dispensable for the integrity of the central area of the coat and is required only to stabilize the coat edges (Antonny and Schekman, 2001).
1.3.2.3. Protein Sorting into COPII vesicles
Export of many secretory proteins from the endoplasmic reticulum (ER) relies on signal-mediated sorting into ER-derived transport vesicles. Experimental evidence demonstrates that certain cargo, in order to be included into COPII vesicles, possess a binding affinity for subunits of the coat. More specifically, pre-budding complexes consisting of Sar1–GTP and Sec23–Sec24 bound to cargo can be isolated under conditions that preserve the Sar–GTP-bound configuration (Kuehn et al. 1998; Aridor et al. 1998). Pre-budding complexes of Sar–Sec23–Sec24–cargo form on the surface of ER membranes. These pre-budding cargo complexes are then gathered by the Sec13–Sec31 complex into nascent vesicles to extract specific cargo from the ER. Presumably, the Sar1 GTPase is regulated in a manner to allow for productive incorporation of pre-budding cargo complexes into the polymerized coat before hydrolysis of bound GTP.
The structure of the pre-budding Sar1–Sec23–Sec24 complex is ‘bowtie-shaped’ and forms a concave surface that apparently faces the ER membrane. Therefore, domains of Sar1 and Sec23–Sec24 could well be available for binding to integral membrane cargo proteins (Bi et al., 2002)
The so called di-acidic sequence (DXE) motif contained within the VSV-G tail sequence is found in many other secretory proteins that are efficiently exported from the ER, including the Kir2.1 potassium channel protein (Ma et al. 2001) and the yeast membrane proteins Sys1p and Gap1p (Malkus et al., 2002; Kappeler et al., 1997). Moreover, Sys1p depends on its di-acidic residues for direct binding to Sec23–Sec24
(Votsmeier and Gallwitz 2001) and Gap1p requires its di-acidic motif to form
pre-budding complexes with Sar1 and Sec23–Sec24 (Malkus et al., 2002) However, there are many other membrane proteins that are efficiently exported from the ER but do not contain apparent di-acidic motifs. For example, the membrane protein ERGIC53, which cycles between the ER and Golgi compartments, has been well studied. This type I transmembrane protein possesses a cytoplasmic tail sequence of 16 residues that is required for properlocalization. More specifically, a conserved pair of aromatic residues at the extreme C-terminus of ERGIC53 is necessary for transport from the
ER (Kappeler et al., 1997). There is some flexibility in this signal as other bulky hydrophobic amino acids can substitute for this C-terminal signal (Nufer et al., 2002). There is also evidence for a role for these terminal residues in binding to the COPII subunits (Kappeler et al., 1997; Nufer et al., 2002). ERGIC53 homologs in yeast also possess bulky hydrophobic residues at their C-termini (LL) that are required for export from the ER and proper localization. Furthermore, when bulky hydrophobic residues are placed at the C-terminus of a transmembrane reporter protein, transport to the Golgi is accelerated (Nufer et al., 2002; Nakamura et al. 1998) although not to rates observed for endogenous ERGIC53.
An additional conserved ER export signal has been identified more recently in the tail sequence of the ERGIC53 family of proteins (Sato and Nakano 2002). This tyrosine-containing motif is ~12 amino acids from the C-terminal signal. Both motifs are required for assembly into COPII pre-budding complexes and for ER export. Other di-aromatic motifs (FF, YY or FY) are found in a similar position in membrane proteins that exit the ER such as the p24 family of proteins (Fiedler et al. 1996; Dominguez et al., 1998) and the Erv41–Erv46 complex (Otte and Barlowe 2002). Interestingly, many of these proteins, including ERGIC53 and VSV-G, form oligomeric complexes, such that a given exported protein would presumably display multiple signals to the COPII budding machinery. Indeed, other reports suggest that multiple signals are needed for efficient export of the Can1p arginine permease (Malkus et al., 2002), the Erv41–Erv46 complex (Otte and Barlowe 2002) and an ATP-binding cassette transporter protein, Yor1p (Epping and Moye-Rowley 2002). A requirement for multiple signals in secretory proteins might be an important element in ER quality control. Recent study has shown that the oligomerization of Emp47p at ER is essential for Emp47p and Emp46p exit from ER (Sato and Nakano 2004).
Several lines of evidence indicate that a family of Sec24 proteins functions in cargo recognition. Furthermore, the presence of multiple Sec24 homologs appears to expand the variety of cargo that must be efficiently exported from the ER. Yeast cells express two additional Sec24-like proteins, termed Lst1 and Iss1. Higher eukaryotes are endowed with at least four Sec24 isoforms (Pagano et al., 1999). In yeast, the Lst1 subunit is not essential for COPII-dependent export but is required for efficient export of specific transmembrane cargoes from the ER (Roberg et al., 1999; Shimoni et al., 2000). Both Sec23–Sec24 and Sec23–Lst1 proteins can be incorporated into a continuous COPII structure, suggesting that heterogeneity in the coat could increase the variety of cargo accommodated by a COPII coated vesicle (Shimoni et al., 2000). More recently, a functional sorting assay demonstrated that both Sec23–Sec24 and
Sec23–Lst1 can function independently in assembly of COPII coats; however, the spectrum of cargo packaged into vesicles synthesized with Sec23–Sec24 was quite distinct from those generated with Sec23–Lst1 (Miller et al., 2002). These observations, coupled with the fact that Sec23–Sec24 displays binding affinities for both di-acidic (Votsmeier and Gallwitz 2001) and di-hydrophobic motifs (Kappeler et al., 1997; Dominguez et al., 1998; Belden and Barlowe 2001), support a direct role for Sec24 in cargo recognition.
Soluble secretory proteins are efficiently exported from the ER and cannot directly contact the COPII coat. Two non-exclusive models, known as the ‘bulk flow’ and ‘receptor-mediated’ export models, have been described in studies addressing export of soluble cargo from the ER. ERGIC53, p24 proteins and Erv29p are proposed to recognize and bind to specific export signals contained within distinct soluble cargo molecules. Presumably, binding is regulated such that fully folded secretory proteins are bound by the receptor in the ER and then released in post-ER compartments. Possible changes in lumenal pH and/or Ca2+ concentration within distinct membrane compartments could regulate receptor–cargo interactions. Alternatively, COPII-dependent oligomerization of membrane receptors could influence receptor–cargo interactions.
1.3.2.4. COPII vesicles in plants
Although the principles of ER-to-GA transport organization in plant cells are supposed to be similar to those in yeast and mammalian systems, evidence in support of such an assertion is largely circumstantial. Moreover, there is a substantial body of evidence that emphasizes the differences, such as the apparent absence of the intermediate compartment in plants, large numbers of GA stacks moving along the ER and the differences in the organization of the cytoskeleton involved in interrelationships between the ER and GA. Although the existence of both anterograde and retrograde ER-to-GA transport in plant cells is generally accepted, no direct evidence about the carriers involved is available. As to COPII vesicles mediating anterograde ER-to-GA transport in yeasts and mammals, they have not been unambiguously identified in plants (Movafeghi et al., 1999; Robinson et al., 1998). However, the existence of Sar1 and other plant homologues of the proteins necessary for their function (d’Enfert et al., 1992; Bar-Peled et al., 1995; Bar-Peled and Raikhel, 1997), as well as microscopical and recent biochemical (Movafeghi et al., 1999) evidence do suggest their existence and a similar role in plants. In this respect, the ability of Sar1 mutants to block ER-to-GA transport in vivo in plant cells
described above is good evidence in support of COPII function in this pathway (Takeuchi et al., 2000).
1.4. SNAREs and Membrane Fusion
Intracellular membrane trafficking in eukaryotic cells utilizes transport vesicles and tubulovesicular structures to deliver cargo proteins and lipids from one internal compartment to the next (Rothman 1994, Salama and Schekman 1995). Membrane fusion is a fundamental biochemical reaction and the final step in all vesicular trafficking events. An ever-widening variety and number of factors are required to mediate the controlled transport of cargo along the secretory pathway. Factors necessary to confer fusion between donor and acceptor compartments (or organelles) include the Rab GTPases, tethering complexes, AAA-type ATPases, and SNAREs (Pfeffer 1999; 2001; Vale 2000). These evolutionarily conserved factors populate the sites of membrane fusion and confer an ordered chain of events that ultimately leads to bilayer fusion. Multiple homologs of these factors are found in eukaryotic cell.
1.4.1. SNAREs
SNAREs were identified after more than a decade of intensive biochemical effort. Using an in vitro trafficking assay developed during the early 1980s, Rothman and colleagues (Balch et al. 1984) were able to purify two soluble proteins required to reconstitute efficient transport. These proteins, N-ethylmaleimide sensitive factor (NSF) and an adaptor protein called soluble NSF attachment protein (SNAP) (Block et al. 1988, Clary et al. 1990), act in many intracellular trafficking pathways. They were subsequently used to affinity purify their membrane receptors from brain, a strategy that yielded proteins crucial for vesicle fusion–mediated neurotransmitter release (Söllner et al. 1993). The receptors were termed SNAREs (for SNAP receptors). Subsequent efforts by a number of groups established that members of the SNARE protein superfamily act not only in neurotransmitter release at the synapse, but also in most, if not every, other intracellular trafficking pathway (Chen & Scheller 2001, Jahn et al. 2003, Kavalali 2002, Pelham 2001, Rizo & Südhof 2002).
The synaptic SNAREs identified by Rothman and colleagues (Söllner et al., 1993) had already been localized either to synaptic vesicles or to the presynaptic membrane (Bennett et al. 1992, Elferink et al. 1989, Südhof et al. 1989). This localization immediately suggested that there were two types of SNAREs, v- (vesicle) SNAREs and t- (target membrane) SNAREs (Söllner et al. 1993). A second nomenclature that
categorizes SNAREs in terms of a single key residue that is usually either arginine (R-SNAREs) or glutamine (Q-(R-SNAREs) has been developed (Fasshauer et al. 1998). Both naming schemes—one functional, the other structural—are still in common use. Because the fusion of two membranes generally requires four SNAREs, at least one of the two membranes must contribute multiple SNAREs. Most intracellular membrane fusion reactions involve one R-SNARE and three Q-SNAREs (Bock et al. 2001, McNew et al. 2000). In many cases, the R-SNARE is contributed by the vesicle, and three Q-SNAREs are contributed by the target organelle.
Assembled SNARE complexes that bridge two membranes are called trans-SNARE complexes. Membrane fusion converts these trans complexes to cis complexes, complexes in which all the SNAREs are associated with the same membrane. The extreme kinetic and thermodynamic stability of cis-SNARE complexes means that their disassembly requires a specialized chaperone machinery (Fasshauer et al. 2002, Fasshauer et al. 1997). This machinery, the chaperone NSF and the cochaperone SNAP, has already been introduced because it was discovered prior to SNAREs and was instrumental in their identification (Söllner et al. 1993a). NSF/SNAP utilizes the energy of ATP hydrolysis to disassemble cis-SNARE complexes, freeing SNAREs for productive trans-complex assembly and allowing the recycling of SNAREs that have already mediated one round of membrane fusion (May et al. 2001).
SNARE proteins are architecturally simple, characterized by the presence of SNARE motifs approximately 60–70 residues in length (Weimbs et al. 1997). In most SNAREs, a single SNARE motif is located immediately adjacent to a C terminal transmembrane anchor. The SNARE motif is most notable for its repeating heptad pattern of hydrophobic residues, spaced such that the adoption of an α-helical structure places all the hydrophobic side chains on the same face of the helix. SNARE motifs assemble into parallel four-helix bundles stabilized by the burial of these hydrophobic helix faces in the bundle core (Antonin et al. 2002b, Poirier et al. 1998, Sutton et al. 1998). The parallel arrangement of SNARE motifs within complexes brings the transmembrane anchors, and the two membranes, into close proximity (Hanson et al. 1997).
In addition to SNARE motifs and membrane anchors, many SNARE proteins have autonomously folding N-terminal domains. The structures of a number of these domains have been determined by using crystallography and/or NMR (Dulubova et al. 2001, Fernandez et al. 1998, Gonzalez et al. 2001, Lerman et al. 2000, Lu et al. 2002, Munson et al. 2000, Tochio et al. 2001). Autonomously folded domains can regulate SNARE assembly. The three-helixbundle domain of syntaxin 7 and the profilin-like
domain of yeast Ykt6p have modest abilities to inhibit SNARE assembly (Antonin et al. 2002a, Tochio et al. 2001). A much more dramatic inhibitory effect is observed for Sso1p. The yeast ortholog of syntaxin, Sso1p, has a three-helix-bundle domain near its N terminus that binds intramolecularly to the SNARE motif. This intramolecular interaction generates a closed conformation that strongly inhibits entry of Sso1p into SNARE complexes, both in vitro and in vivo (Fiebig et al. 1999, Munson et al. 2000, Munson and Hughson 2002).
Fig. 1.5. A general SNARE-mediated fusion reaction. (From Ungar and Hughson 2003)
Most SNAREs are integral membrane proteins with a single transmembrane helix at their C-terminal ends. Within trans-SNARE complexes, at least one of the participating SNAREs is anchored via a transmembrane domain in each membrane.
1.4.2. Other proteins
Other proteins important for vesicle docking and fusion interact either directly or indirectly with SNAREs. Investigators have focused particularly intensively on three families-the rab, tethering, and SM proteins.
1.4.2.1. SM Proteins
A member of the Sec1/Munc18 family (SM) proteins appears to be essential for every intracellular fusion reaction and to act in conjunction with SNAREs (Gallwitz and Jahn 2003).
The Sec1-family of proteins is known to interact with the syntaxin family of SNAREs (Hanson, 2000). This interaction is believed to trigger a change of conformation in the syntaxin (from “closed” to “open”) allowing formation of SNARE complexes (Dulubova et al., 1999). Plants have 6 members of the Sec1 family (Sanderfoot et al., 2000). This number is four-fold lower than the number of syntaxins in Arabidopsis
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